Atomic-Level Characterization of the Chain-Flipping Mechanism in

Letter pubs.acs.org/JPCL

Atomic-Level Characterization of the Chain-Flipping Mechanism in Fatty-Acids Biosynthesis Francesco Colizzi,*,†,§ Matteo Masetti,† Maurizio Recanatini,† and Andrea Cavalli†,‡ †

Department of Pharmacy and Biotechnology, Alma Mater Studiorum-Università di Bologna, via Belmeloro 6, 40126 Bologna, Italy CompuNet, Istituto Italiano di Tecnologia, via Morego 30, 16163 Genova, Italy

‡

S Supporting Information *

ABSTRACT: During fatty acids biosynthesis the elongating acyl chain is sequestered within the core of the highly conserved acyl carrier protein (ACP). At each catalytic step, the acyl intermediates are transiently delivered from ACP to the active site of the enzymatic counterparts and, at the same time, are protected from the solvent to prevent nonselective reactivity. Yet, the molecular determinants of such a universal transitiontermed chain flippingremain poorly understood. Here we capture the atomic-level details of the chainflipping mechanism by using metadynamics simulations. We observe the fatty-acid chain gliding through the protein−protein interface with barely 30% of its surface exposed to water molecules. The small ACP’s helix III acts as gatekeeper of the process, and we find its conformational plasticity critical for a successful substrate transfer. The results are in agreement with a wide range of experimental observations and provide unprecedented insight on the molecular determinants and driving forces of the chain-flipping process.

A

suggesting the connection of this region with the biological function that the carrier protein absolves (Figure 1b,c). Recently, the structural characterization of Escherichia coli acyl-ACP in complex with partner enzymes (FabA4 and LpxD5) has elegantly shown that the acyl chain can flip entirely from the ACP core into the hydrophobic catalytic site of the enzyme. This mechanism, termed chain flipping, appears as a universal strategy3 to access the reactive center of ACP-buried substrates from the catalytic site of enzymatic counterparts.4,5,12,13 Yet, a longstanding and fundamental question remains open: how is such access provided without exposure of the acyl chains to solvent? In this study, we provide the first global view on the entire chain-flipping process and we explicitly address the above question by using advanced molecular dynamics (MD) simulations in explicit solvent. We employed atomistic metadynamics17,18 simulations to map the free-energy landscape of the substrate-delivery process and capture the structural features of relevant free-energy states. Because of its therapeutic potential,19−21 we used the Plasmodium falciparum (Pf) ACP-FabZ model system to investigate the transition of a β-hydroxydecanoyl substrate from the ACP core to the catalytic site of FabZ. The results show that the fatty acid chain can glide through the protein−protein interface in a solvent protected environment. We observed that the protection of the substrate is inherently encoded in the orientation of the α2-α3 ACP region toward the active site of the enzymea feature that is shared among all the ACP-

cyl carrier protein (ACP) is a key element in the biosynthesis of fatty acids and polyketides across all domains of life. ACPs universally sequester, shuttle, and deliver hydrocarbon chains in the cell. In yeast and mammals, ACP is a separate domain of the multifunctional fatty acid synthase polyprotein (type I FAS), whereas it is a small monomeric protein in bacteria and plastids (type II FAS).1 Type II ACPs serve as interaction hub among a series of metabolically related enzymes.2 During fatty acid biosynthesis, the growing acyl substrate is covalently bound to ACP via a flexible prosthetic group (4′ -phosphopantetheine, 4′ -PP) and the acyl chain is buried within ACP’s lipophilic core (Figure 1). This arrangement represents the evolutionary solution to sequester the acyl chain from the solvent and prevent fatty-acid intermediates from nonselective reactivity. Hence, ACP has to recognize several enzymatic counterparts, specifically interact with each of them, and transiently deliver the carried substrate to their active site.3−5 In the last decades, extensive biochemical and structural experiments have been performed to elucidate the structural basis of enzyme recognition and substrate delivery by ACP.3,9,10 ACP-enzyme interactions are mostly driven by the electrostatic complementarity between the conserved acidic residues on α2 and α3 ACP helices and the arginine/lysine-rich region surrounding the active site of ACP-dependent enzymes. Additionally, specific lipophilic patches account for selectivity among different ACP-interacting partners.3 NMR data and crystal structure B-factors have shown that ACP’s α3-helix is the most dynamic segment of the protein and its motion seems determinant for the release of the substrate upon enzyme interaction.3,11 Clusters of evolutionarily conserved residues are present around the cleft between α2 and α3 ACP helices, © XXXX American Chemical Society

Received: June 6, 2016 Accepted: July 13, 2016

2899

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904

Letter

The Journal of Physical Chemistry Letters

generated, that manifested the typical funnel-like energy landscape associated with the native state of a protein−protein complex.16 Hence we selected this binding mode for further investigation. The docked ACP-FabZ interface was formed by ACP helices α2 and α3 (and the loop connecting them) whose acidic residues nicely complemented the positive patch surrounding the active site of FabZ. X-ray crystallography,12,13,22 chemical shift perturbation,4,23 site-directed fluorescence analyses,24 and computational25 studies have identified the α2-α3 region as component of the ACP-enzyme interacting interface.22−25 In agreement with the biochemical and structural observations of Nguyen et al.,4 we found that the docking interface of Pf FabZ is different from the one observed in the E. coli Acp-FabA complex, thus reflecting the shift of the “positive patch” of FabZ enzymes when compared to FabA ones (see Supporting Information, SI).4 To gain an atomistic description of the chain-flipping mechanism, we then used the docked ACP-FabZ complex to study the transition of a β-hydroxydecanoyl substrate from the ACP core to the active site of FabZ (Figure 3). The configurational space was explored with metadynamics, an enhanced-sampling method that allows one to efficiently reach biologically relevant time scales (see Methods).26,27 We observed that the substrate was constantly protected from the solvent throughout the entire chain-flipping process (Figure 3a, upper panel). The solvation of the β-hydroxydecanoyl moiety reached only ∼30% of its maximum (defined as the value corresponding to moiety freely diffusing in water) in correspondence of a metastable region of the free-energy landscape (Figure 3a, lower panel). Furthermore, the low solvation was independent from the exact pathway followed by the substrate (i.e., independent from the distance f rom the path in Figure 3a). Thus, we argue that the protection of the reactive center from the solvent does not strictly depend on the chainflipping path but it is rather encoded in the orientation of the α2 and α3 helices of ACP toward the active site of the enzyme. Such an orientation is a common feature among all the characterized ACP-enzyme complexes,4,5,12,13 and it makes the substrate-delivery process tolerant to binding mode variability. In this respect, although the ACP-enzyme complexes show different binding modes,4,5,12,13 the conserved orientation of the α2−α3 ACP region toward the active site warranties the protection of the substrate from the solvent during reiterated cycles of chain elongation. The carried and delivered states of the substrate belonged to similarand well-definedfree-energy basins separated by a wide metastable region (underlying an intermediate state) and a main transition state (TS, Figure 3). The egress of the substrate was driven by the extrusion of the hydrocarbon chain through the cleft lined by hydrophobic residues between ACP helices α2 and α3 (Figure 3b,c). During the egress process, the small α3 exhibited large conformational fluctuations coupled to the transient flooding of ACP core. Accordingly, extensive movements of helix α3 during chain flipping have been postulated on the basis of chemical shift measurements and the crystal structure of a cross-linked ACP-FabA complex in E. coli.4 In particular, it has been argued that some transient destabilization of the ACP hydrophobic core upon interaction with cognate enzymes might allow the substrate to partition into the hydrophobic active site channel of the enzyme.3 The short helix α3 has a high density of charged residues (Figure 1c) and has been experimentally observed in a helix-loop conformational equilibrium by solution-NMR (Figure 4a);28 it

Figure 1. Structure and sequence of the acyl carrier protein (ACP). (a) Overall fold of Pf ACP loaded with a β-hydroxydecanoyl substrate (carbon atoms in cyan stick); the α3 helix (residues 57−62) is show in yellow ribbons while other helical structures are in violet. A slice of the molecular surface at the level of the acyl binding pocket (that is shaped by a bundle of α helices) is shown in blue. (b) The molecular surface of Pf ACP mapped with conservation scores based on phylogenetic relations between homologous sequences.6,7 Green spots represent highly conserved regions suggesting a functional role on the ACP surface. (c) Sequence alignment of ACP homologues with identical residues in green. Alignment generated using the structural matrix implemented in BODIL.8

protein complexes characterized so far. Moreover, we identify the small helix III in ACP as gatekeeper of the transfer mechanism, and we quantify the role of its structural plasticity in the delivery process. Our results complement with atomic spatiotemporal resolution the most recent experimental insight and offer a structural interpretation to the strategy evolved for preventing cross-reactivity during fatty acid biosynthesis. We found that the ACP-FabZ complex shown in Figure 2 was the only configuration, among the top-ranked poses

Figure 2. Interaction model for the ACP-FabZ complex in Pf FabZ and ACP structures are represented in green and white ribbons, respectively. The ACP’s α2 (in violet) and α3 (in yellow) helices at the protein−protein interface are highlighted. The molecular surfaces are colored by the electrostatic potential showing the complementarity between the docked interfaces; contours are at +8 (in blue) and −8 (in red) kBT/e. The corresponding energetic binding funnel obtained from the refinement with RosettaDock14,15 is shown in the bottom right corner. The presence of a stable energy minimum surrounded by a broad region of attraction (funnel) is associated with the native state of a protein−protein complex.16 2900

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904

Letter

The Journal of Physical Chemistry Letters

Figure 3. Atomistic and energetic features of the substrate delivery process in the ACP-FabZ complex. (a) Free-energy landscape of the delivery process reconstructed as a function of the progress along, and the distance from, the reference path. The encircled numbers correspond to the carried, metastable, and delivered states shown in panel c. The upper panel shows the % of solvation of the β-hydroxydecanoyl moiety with respect to the fully solvated one; the maximum solvation is between steps 8−10. (b) Graphical representation of the chain-flipping process obtained from the simulations; the range of color (from red to blue for the carried and delivered state, respectively) represents the progression along the path. (c) Snapshots of the metadynamics trajectory showing the β-hydroxydecanoyl substrate (carbon atoms in cyan) embedded into the hydrophobic core of ACP (1), at the interface between ACP and FabZ (2), and delivered to the active site of FabZ (3). The α2 and α3 helices of ACP are shown in violet and yellow ribbons, respectively; FabZ structure is in transparent green ribbons and the catalytic residues, His 133′ and Glu147, are in sticks (carbon atoms in cyan).

substrate release from ACP core doubled in comparison to the unrestrained simulations (Figure 4b). Hence, the plasticity of the short α3 helix is a key element in facilitating the release of substrate at the ACP−FabZ interface. The helix α3 has been observed at the protein−protein interface of all the ACP− enzyme complexes reported so far.4,5,12,13 We argue that its plasticity is a universal feature required for the chain-flipping process. After the release from ACP, the substrate populated the ACP−FabZ interface with only its acyl moiety protruding toward FabZ active site (intermediate state shown in Figure 3). This intermediate state facilitated the accumulation of the substrate at ACP−FabZ interface in proximity of the transition state. We identified the TS as the event in which the reactive center of the substrate actually flips from ACP to FabZ. Yet, the characterization of the TS is out of the aims of this work, and it is only briefly detailed in the SI, Figure S3. Within the binding site of FabZ, the substrate was driven into a catalytically competent configuration by a broad free-energy basin of ligand attraction (Figure 5). We found a stable precatalytic binding mode involving the H-bond between the backbone nitrogen of Phe 171 and the thioester carbonyl group of the substrate (Figure 5a). The observed binding mode showed high similarity with respect to the crystal structure of mechanism-based probes cross-linked to bacterial FabA (Figure 5a, left panel).4,29 Thus, the overlay of the MD and crystallographic structures directly suggest the putative gateway for substrate access or product release from the active site. The precatalytic pose was further stabilized by the interaction of the

Figure 4. Dynamics of ACP’s α3 helix in the delivery of substrate. (a) Sausage representation of Pf ACP structure from a solution-NMR ensemble.28 The width of the tube matches the width of the ensemble and highlights the inherent conformational variability of helix α3 and of the adjacent loop-connection to helix α2. (b) Free-energy profile of the early stages of the delivery process as a function of the path s. When the α3 helix is restrained to its helical conformation (light gray plot), the energy barrier is higher than the equivalent unrestrained simulation (dark gray plot).

is likely that such a helix-loop equilibrium is perturbed, or conformationally selected, by the positive electrostatic patch of partner enzymes. In this scenario, the transient flooding of the ACP core would drive the progression of the acyl substrate toward the hydrophobic active site of FabZ. To assess the effect of ACP structural plasticity on the delivery process, we artificially restrained the α3 conformation to its native α-helical geometry. Strikingly, when preventing α3 conformational freedom, the free-energy barrier for the 2901

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904

Letter

The Journal of Physical Chemistry Letters

In conclusion, the atomistic simulations presented herein show the flipping of the substrate through an ACP-enzyme interface formed by helices α2 and α3 of ACP. The concurrent protection of the acyl substrate from the solventand thus the reduction of nonselective cross reactivityis intrinsically encoded in the orientation of α2 and α3 ACP helices toward the active site of the enzyme. We assessed the role of the conformational freedom of ACP helix α3 in the release of the substrate from the carrier, and we argue that its plasticity is a universal feature required for the chain-flipping process. The progression of the substrate along the delivery pathway is facilitated by a destabilization of the ACP core and a widely populated intermediate state. Furthermore, a deep basin of ligand attraction favors the arrangement of the active site into a precatalytic configuration that drives the system toward the catalytically competent configuration. The results presented in this report provide an unprecedented atomic-level characterization of the chain-flipping process and offer a mechanicistic reference to further the molecular understanding of ACP-based metabolic pathways.

■

METHODS Protein−Protein Docking. A two-step docking strategy was used to generate the converged ACP-FabZ mode of interaction shown in Figure 2 (procedure detailed in the SI). Briefly, after a global configurational search driven by shape-complementarity criteria,31−33 top-ranked docking poses were locally refined with a detailed energy function and accounting for side-chain flexibility.14,15 The local refinement allows the detection of binding poses that are surrounded by a broad region of attraction (funnel) on the energy surface. The pose showing a funnel-like energy landscape is typically associated with the native state.16 ACP-enzyme docking techniques have been successfully used elsewhere for enzyme engineering.34 Metadynamics Simulations. Metadynamics is a well-established enhanced-sampling method that allows one to efficiently reach biologically relevant time scales with MD and to reconstruct the underlying free-energy of the process under investigation.26,27 Here, the underlying free energy was reconstructed as a function of two collective variables (CVs), or reaction coordinates, based on a preassigned path built on previous experimental insight.4,29,30 The initial and final states of the path corresponded to the substrate embedded into the ACP core (referred to as “carried” state) and into the active site of FabZ (“delivered” state), respectively. In this framework, the microscopic coordinates of the system, q, are mapped in the CV space by s(q), that measures the progress along the path, and z(q) that measures the distance from the preassigned path.35 Using these variables, one can explore the free-energy landscape between an initial and final state and can find low free-energy pathways connecting thempathways that in turn can be different from the originally assigned one.35 This approach has been successfully used to study configurational changes in a variety of complex (bio)molecular systems.36−40 The methodology employed to parametrize the path connecting the carried and delivered states of the β-hydroxydecanoyl substrate is in the SI.

Figure 5. Dynamics of the substrate within FabZ active site. (a) Representative system configurations (carbon atoms of the substrate are in cyan and those of FabZ in green) and underlying free-energy profile as a function of the distance from the path in the delivered state. The X-ray crystallography binding mode (carbon atoms in white, pdb code: 1mka29) of a mechanism-based probe overlays with the precatalytic configuration disclosed by the MD simulations. (b) Schematic representation of the mechanism of substrate dehydration reproduced in the catalytically competent configuration of panel a. The N-terminal positive dipole moment of FabZ helix α2 (green ribbon) further stabilizes the interaction of the thioester carbonyl group of the ACP-bound substrate and the backbone of Gly 152. The enzymatic mechanism is based on previous structural insight.4,29,30

β-hydroxy group of the substrate with the side chain of Glu 147. The flipping of the thioester carbonyl toward the nitrogen of Gly 141 and the rearrangement of water molecules around the active site allowed a close proximity between the Nϵ2 of catalytic His 133′ and the polarized α-C−H bond of the substrate (Figure 5a, 5b). This configuration is thought to trigger the chemical reactivity of the substrate-enzyme complex and was stereochemically consistent with the formation of the natural trans-2-decenoyl product as suggested by previous structural studies.4,29,30 It still remains an open question how the product of ACPdependent enzymatic reactions can return to the ACP cavity to permit further modifications of the acyl chain along the metabolic pathway.3 We argue that the features of the reaction product as well as those of the transition state (see Figure S3 and related text) contribute in modulating the release of the product into the ACP core. We hope this question will be the focus of future investigations.

■

ASSOCIATED CONTENT

* Supporting Information S

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.jpclett.6b01230. 2902

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904

Letter

The Journal of Physical Chemistry Letters

■

complex required for biotin synthesis. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 17406−17411. (13) Cryle, M. J.; Schlichting, I. Structural insights from a P450 Carrier Protein complex reveal how specificity is achieved in the P450BioI ACP complex. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 15696−15701. (14) Lyskov, S.; Gray, J. J. The RosettaDock server for local proteinprotein docking. Nucleic Acids Res. 2008, 36, W233−W238. (15) Gray, J. J.; Moughon, S.; Wang, C.; Schueler-Furman, O.; Kuhlman, B.; Rohl, C. A.; Baker, D. Protein-protein docking with simultaneous optimization of rigid-body displacement and side-chain conformations. J. Mol. Biol. 2003, 331, 281−299. (16) Kozakov, D.; Schueler-Furman, O.; Vajda, S. Discrimination of near-native structures in protein-protein docking by testing the stability of local minima. Proteins: Struct., Funct., Genet. 2008, 72, 993−1004. (17) Laio, A.; Parrinello, M. Escaping free-energy minima. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 12562−12566. (18) Bussi, G.; Laio, A.; Parrinello, M. Equilibrium free energies from nonequilibrium metadynamics. Phys. Rev. Lett. 2006, 96, 090601. (19) Shears, M. J.; Botté, C. Y.; McFadden, G. I. Fatty acid metabolism in the Plasmodium apicoplast: Drugs, doubts and knockouts. Mol. Biochem. Parasitol. 2015, 199, 34−50. (20) van Schaijk, B. C. L.; Kumar, T. R. S.; Vos, M. W.; Richman, A.; van Gemert, G.-J.; Li, T.; Eappen, A. G.; Williamson, K. C.; Morahan, B. J.; Fishbaugher, M.; et al. Type II Fatty Acid Biosynthesis Is Essential for Plasmodium falciparum Sporozoite Development in the Midgut of Anopheles Mosquitoes. Eukaryotic Cell 2014, 13, 550−559. (21) Vaughan, A. M.; O’Neill, M. T.; Tarun, A. S.; Camargo, N.; Phuong, T. M.; Aly, A. S.; Cowman, A. F.; Kappe, S. H. Type II fatty acid synthesis is essential only for malaria parasite late liver stage development. Cell. Microbiol. 2009, 11, 506−520. (22) Parris, K. D.; Lin, L.; Tam, A.; Mathew, R.; Hixon, J.; Stahl, M.; Fritz, C. C.; Seehra, J.; Somers, W. S. Crystal structures of substrate binding to Bacillus subtilis holo-(acyl carrier protein) synthase reveal a novel trimeric arrangement of molecules resulting in three active sites. Structure 2000, 8, 883−895. (23) Zhang, Y.-M.; Wu, B.; Zheng, J.; Rock, C. O. Key residues responsible for acyl carrier protein and β-ketoacyl-acyl carrier protein reductase (FabG) interaction. J. Biol. Chem. 2003, 278, 52935−52943. (24) Worsham, L. M.; Earls, L.; Jolly, C.; Langston, K. G.; Trent, M. S.; Ernst-Fonberg, M. L. Amino acid residues of Escherichia coli acyl carrier protein involved in heterologous protein interactions. Biochemistry 2003, 42, 167−176. (25) Zhang, Y.-M.; Marrakchi, H.; White, S. W.; Rock, C. O. The application of computational methods to explore the diversity and structure of bacterial fatty acid synthase. J. Lipid Res. 2003, 44, 1−10. (26) Cavalli, A.; Spitaleri, A.; Saladino, G.; Gervasio, F. L. Investigating Drug−Target Association and Dissociation Mechanisms Using Metadynamics-Based Algorithms. Acc. Chem. Res. 2015, 48, 277−285. (27) Bussi, G.; Branduardi, D. Free-Energy Calculations with Metadynamics: Theory and Practice. Reviews in Computational Chemistry Volume 28 2015, 1−49. (28) Sharma, A. K.; Sharma, S. K.; Surolia, A.; Surolia, N.; Sarma, S. P. Solution structures of conformationally equilibrium forms of holoacyl carrier protein (PfACP) from Plasmodium falciparum provides insight into the mechanism of activation of ACPs. Biochemistry 2006, 45, 6904−6916. (29) Leesong, M.; Henderson, B. S.; Gillig, J. R.; Schwab, J. M.; Smith, J. L. Structure of a dehydratase−isomerase from the bacterial pathway for biosynthesis of unsaturated fatty acids: two catalytic activities in one active site. Structure 1996, 4, 253−264. (30) Kostrewa, D.; Winkler, F. K.; Folkers, G.; Scapozza, L.; Perozzo, R. The crystal structure of PfFabZ, the unique β-hydroxyacyl-ACP dehydratase involved in fatty acid biosynthesis of Plasmodium falciparum. Protein Sci. 2005, 14, 1570−1580.

Supplementary methods and supplementary discussion (PDF) Animation of the chain-flipping trajectory (AVI)

AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Present Address §

(F.C.) Institute for Research in Biomedicine (IRB Barcelona) Carrer Baldiri Reixac 10, 08028, Barcelona, Spain Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS We acknowledge the CINECA (Casalecchio di Reno, Italy) for the availability of high performance computing resources and support. The authors thank Ivan Ivani at IRB Barcelona for reading the manuscript and providing insightful feedback. F.C. thanks Laurène Bastet, Gaston Giroux, and the Bibliothèque Roger-Maltais at the Université de Sherbrooke in Québec, Canada for providing infrastructures and support. F.C. acknowledges Sabbatical Funding from Romano Colizzi & Maria Gaudio in Taranto, Italy.

■

REFERENCES

(1) Byers, D. M.; Gong, H. Acyl carrier protein: structure-function relationships in a conserved multifunctional protein family. Biochem. Cell Biol. 2007, 85, 649−662. (2) White, S. W.; Zheng, J.; Zhang, Y.-M.; Rock, C. O. The structural biology of type II fatty acid biosynthesis. Annu. Rev. Biochem. 2005, 74, 791−831. (3) Cronan, J. E. The chain-flipping mechanism of ACP (acyl carrier protein)-dependent enzymes appears universal. Biochem. J. 2014, 460, 157−163. (4) Nguyen, C.; Haushalter, R. W.; Lee, D. J.; Markwick, P. R. L.; Bruegger, J.; Caldara-Festin, G.; Finzel, K.; Jackson, D. R.; Ishikawa, F.; O’Dowd, B.; et al. Trapping the dynamic acyl carrier protein in fatty acid biosynthesis. Nature 2014, 505, 427−431. (5) Masoudi, A.; Raetz, C. R.; Zhou, P.; Pemble, C. W., IV Chasing acyl carrier protein through a catalytic cycle of lipid A production. Nature 2014, 505, 422−426. (6) Glaser, F.; Pupko, T.; Paz, I.; Bell, R. E.; Bechor-Shental, D.; Martz, E.; Ben-Tal, N. ConSurf: identification of functional regions in proteins by surface-mapping of phylogenetic information. Bioinformatics 2003, 19, 163−164. (7) Celniker, G.; Nimrod, G.; Ashkenazy, H.; Glaser, F.; Martz, E.; Mayrose, I.; Pupko, T.; Ben-Tal, N. ConSurf: using evolutionary data to raise testable hypotheses about protein function. Isr. J. Chem. 2013, 53, 199−206. (8) Lehtonen, J. V.; Still, D.-J.; Rantanen, V.-V.; Ekholm, J.; Björklund, D.; Iftikhar, Z.; Huhtala, M.; Repo, S.; Jussila, A.; Jaakkola, J.; et al. BODIL: a molecular modeling environment for structure-function analysis and drug design. J. Comput.-Aided Mol. Des. 2004, 18, 401−419. (9) Chan, D.; Vogel, H. Current understanding of fatty acid biosynthesis and the acyl carrier protein. Biochem. J. 2010, 430, 1−19. (10) Crosby, J.; Crump, M. P. The structural role of the carrier protein-active controller or passive carrier. Nat. Prod. Rep. 2012, 29, 1111−1137. (11) Colizzi, F.; Recanatini, M.; Cavalli, A. Mechanical features of Plasmodium falciparum acyl carrier protein in the delivery of substrates. J. Chem. Inf. Model. 2008, 48, 2289−2293. (12) Agarwal, V.; Lin, S.; Lukk, T.; Nair, S. K.; Cronan, J. E. Structure of the enzyme-acyl carrier protein (ACP) substrate gatekeeper 2903

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904

Letter

The Journal of Physical Chemistry Letters (31) Duhovny, D.; Nussinov, R.; Wolfson, H. J. Efficient Unbound Docking of Rigid Molecules. In Algorithms in Bioinformatics; Springer: Berlin, 2002; pp 185−200. (32) Schneidman-Duhovny, D.; Inbar, Y.; Nussinov, R.; Wolfson, H. J. PatchDock and SymmDock: servers for rigid and symmetric docking. Nucleic Acids Res. 2005, 33, W363−W367. (33) Mashiach, E.; Schneidman-Duhovny, D.; Peri, A.; Shavit, Y.; Nussinov, R.; Wolfson, H. J. An integrated suite of fast docking algorithms. Proteins: Struct., Funct., Genet. 2010, 78, 3197−3204. (34) Jiménez-Osés, G.; Osuna, S.; Gao, X.; Sawaya, M. R.; Gilson, L.; Collier, S. J.; Huisman, G. W.; Yeates, T. O.; Tang, Y.; Houk, K. N. The role of distant mutations and allosteric regulation on LovD active site dynamics. Nat. Chem. Biol. 2014, 10, 431−436. (35) Branduardi, D.; Gervasio, F. L.; Parrinello, M. From A to B in free energy space. J. Chem. Phys. 2007, 126, 054103. (36) Berteotti, A.; Cavalli, A.; Branduardi, D.; Gervasio, F. L.; Recanatini, M.; Parrinello, M. Protein conformational transitions: the closure mechanism of a kinase explored by atomistic simulations. J. Am. Chem. Soc. 2009, 131, 244−250. (37) Pfaendtner, J.; Branduardi, D.; Parrinello, M.; Pollard, T. D.; Voth, G. A. Nucleotide-dependent conformational states of actin. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 12723−12728. (38) Ceccarini, L.; Masetti, M.; Cavalli, A.; Recanatini, M. Ion conduction through the hERG potassium channel. PLoS One 2012, 7, e49017. (39) Franco, D.; Vargiu, A. V.; Magistrato, A. Ru [(bpy) 2 (dppz)] 2+ and rh [(bpy) 2 (chrysi)] 3+ targeting double strand DNA: The shape of the intercalating ligand tunes the free energy landscape of deintercalation. Inorg. Chem. 2014, 53, 7999−8008. (40) Formoso, E.; Limongelli, V.; Parrinello, M. Energetics and structural characterization of the large-scale functional motion of adenylate kinase. Sci. Rep. 2015, 5, 8425.

2904

DOI: 10.1021/acs.jpclett.6b01230 J. Phys. Chem. Lett. 2016, 7, 2899−2904